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. Author manuscript; available in PMC: 2015 Jul 3.
Published in final edited form as: Mol Cell. 2014 May 29;55(1):31–46. doi: 10.1016/j.molcel.2014.04.028

HDAC6 Deacetylates and Ubiquitinates MSH2 to Maintain Proper Levels of MutSα

Mu Zhang 1,15, Shengyan Xiang 1,15, Heui-Yun Joo 2, Lei Wang 1, Kendra A Williams 1, Wei Liu 1, Chen Hu 1, Dan Tong 3, Joshua Haakenson 1, Chuangui Wang 4, Shengping Zhang 4, Ryan E Pavlovicz 5, Amanda Jones 2, Kristina H Schmidt 6,7, Jinfu Tang 1, Huiqin Dong 1, Bin Shan 8, Bin Fang 9, Rangasudhagar Radhakrishnan 7, Peter M Glazer 10, Patrick Matthias 11, John Koomen 9, Edward Seto 7, Gerold Bepler 12, Santo V Nicosia 1,13, Jiandong Chen 7, Chenglong Li 5,14, Liya Gu 3, Guo-Min Li 3, Wenlong Bai 1,7, Hengbin Wang 2, Xiaohong Zhang 1,7,*
PMCID: PMC4188514  NIHMSID: NIHMS629308  PMID: 24882211

SUMMARY

MutS protein homolog 2 (MSH2) is a key DNA mismatch repair protein. It forms the MSH2-MSH6 (MutSα) and MSH2-MSH3 (MutSβ) heterodimers, which help to ensure genomic integrity. MutSα not only recognizes and repairs mismatched nucleotides but also recognizes DNA adducts induced by DNA-damaging agents, and triggers cell-cycle arrest and apoptosis. Loss or depletion of MutSα from cells leads to microsatellite instability (MSI) and resistance to DNA damage. Although the level of MutSα can be reduced by the ubiquitin-proteasome pathway, the detailed mechanisms of this regulation remain elusive. Here we report that histone deacetylase 6 (HDAC6) sequentially deacetylates and ubiquitinates MSH2, leading to MSH2 degradation. In addition, HDAC6 significantly reduces cellular sensitivity to DNA-damaging agents and decreases cellular DNA mismatch repair activities by downregulation of MSH2. Overall, these findings reveal a mechanism by which proper levels of MutSα are maintained.

INTRODUCTION

HDAC6 was first cloned from mouse and human as a mammalian homolog of yeast HDA1 (Grozinger et al., 1999; Verdel and Khochbin, 1999). Uniquely, HDAC6 contains two functional tandem deacetylase domains, termed DAC1 and DAC2, as well as a ZnF-UBP domain, which is a zinc finger-containing region that is homologous with the noncatalytic domain of several ubiquitin-specific proteases (USPs) (Seigneurin-Berny et al., 2001). The HDAC6 ZnF-UBP domain is capable of binding to mono- or polyubiquitin, as well as ubiquitinated proteins (Boyault et al., 2006; Hook et al., 2002; Seigneurin-Berny et al., 2001). HDAC6’s substrates include cytosolic proteins, such as α-tubulin, HSP90, and cortactin. (Hubbert et al., 2002; Kovacs et al., 2005; Zhang et al., 2007). HDAC6 acts in ubiquitin-dependent autophagy by allowing the processing or degradation of protein aggregates (Pandey et al., 2007). Additionally, HDAC6 is involved in misfolded protein-induced cell stress (Kawaguchi et al., 2003). HDAC6 is now considered as a master regulator of the cellular response to cytotoxic assaults (Matthias et al., 2008). We and others recently reported that HDAC6 plays a role in genotoxic stress response (Namdar et al., 2010; Wang et al., 2012); however, the underlying mechanisms are unclear.

MSH2 is an essential component in eukaryotic DNA mismatch repair (MMR), a major genome maintenance system that ensures genetic stability by correcting DNA biosynthetic errors, suppressing non-homologous recombination, and mediating DNA damage signaling (Li, 2008). As an obligate subunit for mismatch recognition proteins in eukaryotic cells, MSH2 interacts with MSH6 or MSH3 to form the MutSα or MutSβ complexes, respectively. MutSα specifically recognizes single base-base mismatches and 1–2 nucleotide insertion/deletion mispairs, whereas MutSβ recognizes large insertion/deletion heteroduplexes (Drummond et al., 1995; Genschel et al., 1998). Recently, the human MMR reaction has been reconstituted using purified proteins (Constantin et al., 2005; Zhang et al., 2005). It is well accepted that MMR is initiated by binding of MutSα or MutSβ to a DNA mispair. This reaction triggers concerted interactions among MutSα, MutLα (MLH1-PMS2), proliferating cellular nuclear antigen (PCNA), and replication protein A (RPA), facilitating communications between the mismatch and a strand break and leading to recruitment of exonuclease 1 (EXO1) to the strand break. EXO1 then excises nascent DNA from the nick toward and beyond the mismatch to generate a single-strand gap, which is filled by polymerase δ using the parental DNA strand as template. Finally, the nick is ligated by DNA ligase I. The importance of MMR in genome maintenance is underscored by the fact that defects in key MMR genes, such as MSH2 and MLH1, lead to genome-wide instability and predisposition to cancers (Kolodner, 1995; Li, 2008; Modrich, 1994). In addition to correcting mismatches, the genome maintenance function of the MMR system includes mediating DNA damage-induced apoptosis. MutSα plays an important role in this process by specifically recognizing DNA lesions caused by many DNA-damaging agents, including 6-thioguanine (6-TG), N-methyl- N′-nitro-N-nitrosoguanidine (MNNG), cisplatin, carboplatin, doxorubicin, and etoposide (Fink et al., 1998). However, this recognition fails to remove these DNA adducts from DNA; instead, it triggers a futile repair cycle (Li, 1999), thereby leading to apoptosis. In contrast, cells lacking MutSα or expressing a low level of MutSα are highly resistant to the treatment of DNA-damaging agents (Li, 2008). Thus, MutSα serves as a DNA damage sensor.

Here we report that the levels of MutSα are controlled through MSH2 acetylation in vivo. HDAC6 interacts with MSH2 and regulates MSH2’s turnover via sequential deacetylation and ubiquitination. Interestingly, we have uncovered a novel ubiquitin E3 ligase activity of HDAC6, for which its DAC1 domain is responsible, while the deacetylase activity of HDAC6 mainly resides in its DAC2 domain. HDAC6 confers cellular resistance to 6-TG and MNNG and reduces cellular DNA MMR activities by downregulation of MSH2. Taken together, our findings provide a novel mechanism by which HDAC6 regulates genotoxic stress response and MMR activities.

RESULTS

HDAC6 Interacts with MSH2 In Vivo

To investigate the nuclear function of HDAC6, we set out to identify proteins associated with HDAC6 in HeLaS3 nuclear extracts (NEs). We first fractionated NEs onto a phosphocellulose P11 column (Figure 1A) and HDAC6 was exclusively distributed in BC300 (Figure S1A). The BC300 fraction was further separated through a DEAE5PW column and HDAC6 was eluted between 133–270mMKCl. We then used affinity chromatography to identify proteins associated with HDAC6. The unique bands existing in the anti-HDAC6, but not in the control anti-IgG immunoprecipitates, were excised and analyzed by liquid chromatography-tandem mass spectrometry (LC-MS/MS) (Figure 1B). The mass spectrometry analysis identified four DNA MMR proteins, MSH2, MSH3, MSH6 and MLH1, as well as two proteins involved in DNA double stranded break repair, MRE11 and RAD50, in the HDAC6 nuclear complex (Table S1). To confirm that these proteins indeed form a complex, we fractionated the DEAE5PW pool through a Superose 6 gel filtration column. As shown in Figure 1C, HDAC6, MSH2, MSH6, MLH1, MSH3, MRE11 and RAD50 were co-eluted in fraction 33, indicating that HDAC6 may form a ~2,000 KD complex. To investigate whether HDAC6 interacts with MSH2 in non-transformed cell lines, the NEs from T29, a non-transformed ovarian surface epithelial cell line, were fractionated by a Superose 6 gel filtration column. As shown in Figure 1D, HDAC6 is co-purified with MSH2 in fraction 34, indicating that HDAC6 may form a 2,000 KD nuclear complex with MSH2 in T29 cells as it does in HeLa cells. Since MSH2 is a key DNA MMR protein, we then focused on verifying the HDAC6 and MSH2 interaction by employing coimmunoprecipitation (coIP) assays. As shown in Figure 1E, anti-HDAC6 antibodies (lane 3), but not anti-IgG control antibodies (lane 2), immunoprecipitated MSH2 in HeLa cell extracts. In a reciprocal fashion, anti-MSH2 antibodies specifically immunoprecipitated HDAC6 in 293T cells (Figure 1F). Our results confirmed that endogenous MSH2 and HDAC6 interact in vivo.

Figure 1. HDAC6 Interacts with MSH2 In Vivo.

Figure 1

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(A) The scheme of the steps used to identify HDAC6-interacting proteins.

(B) The 133–270mMfraction was immunoprecipitated (IPed) with either anti-IgG or anti-HDAC6 antibody followed by silver stain. M refers to the molecular weight marker.

(C) Superose 6 fractions were subjected to western blotting (WB) analyses as indicated.

(D) HDAC6 was copurified in T29 cell NEs fractioned by Superose 6 gel filtration column. The indicated fractions from the column were subjected to TCA (trichloroacetic acid) precipitation, and the WB analyses were carried out with indicated antibodies.

(E) HeLa cell extracts were IPed with either an anti-IgG or an anti-HDAC6 antibody. The WB analyses were performed as indicated.

(F) The reciprocal IP-WBs were performed as indicated. See also Figures S1 and S2 and Table S1.

We next tested which domain of HDAC6 binds to MSH2 and vice versa. In cotransfection experiments, we found that the N-terminal region of HDAC6 (1–503), encompassing the DAC1 domain, binds to MSH2 (Figures S1B and S1C). Human MSH2 protein can be divided into five domains according to its crystal structure, namely mismatch binding, connector, lever, clamp, and ATPase domains (Warren et al., 2007). GST bead-bound MSH2 full-length and MSH2 deletion mutants purified from bacteria were used to pull down Flag-HDAC6 in Sf9 cells transduced with Flag-HDAC6 viruses. As shown in Figures S1D and S1E, two MSH2 regions: part of the lever domain (533–624) and the ATPase domain (624–700) are essential for MSH2 binding to HDAC6. In addition, we found that HDAC6 binds to MSH6, MLH1, and NBS1 (Figures S2A–S2C).

HDAC6, Not SirT2, Deacetylates MSH2

Because HDAC6 is a deacetylase, we first examined whether HDAC6 deacetylates MSH2. As shown in Figure 2A, overexpression of F-HDAC6 in 293T cells drastically reduced the level of acetylated MSH2. In contrast, acetylated MSH2 was elevated in HDAC6 knockout (KO) mouse embryonic fibroblasts (MEFs) and HDAC6 knockdown (KD) C13 cells (Figures 2B and 2C) as compared with their counterparts, further indicating that HDAC6 is an MSH2 deacetylase in vivo. Moreover, several pharmacological HDAC inhibitors were used to examine whether the level of acetylated MSH2 inversely correlates with HDAC6 deacetylase activity. As shown in Figure 2D, the level of acetylated MSH2 was increased after tubacin (an HDAC6-selective inhibitor) treatment compared with the vehicle control, but not after sodium butyrate (NaB, a pan-HDAC inhibitor, which does not suppress HDAC6 activity efficiently) treatment. In addition, SAHA, a clinically relevant HDAC inhibitor, increased MSH2 acetylation (Figure 2E). Previously, a class III HDAC, SirT2, was shown to deacetylate α-tubulin (North et al., 2003), which is an HDAC6 substrate (Hubbert et al., 2002). We therefore examined whether SirT2 deacetylates MSH2 as well. As shown in Figures 2F and 2G, neither the SirT2 inhibitor, AGK2, nor the siRNA of SirT2 could notably induce MSH2 acetylation. Therefore, HDAC6, not SirT2, is the major MSH2 deacetylase in vivo. In addition, HDAC6 may deacetylate MSH6 and MLH1 (Figures S2D and S2E).

Figure 2. HDAC6 Deacetylates MSH2.

Figure 2

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(A) 293T cells were transfected with either HA-MSH2 alone or HA-MSH2 with F-HDAC6. IP-WB and direct WB were performed as indicated.

(B) The levels of acetylated MSH2 are enhanced in HDAC6 knockout (KO) MEFs. IP-WB and direct WB were performed as indicated. Please note that in WT and KO MEFs, equal amounts of immunoprecipitated MSH2, not equal amounts of cell lysates, were loaded on SDS-PAGE. mHD6 represents mouse HDAC6.

(C) C13 cells were stably transfected with control shRNA(pRS) and HDAC6shRNA (Origene) to generate control (C13-pRS) and HDAC6 knockdown cells (C13-HD6KD), respectively. IP-WB and direct WB were performed as indicated. Please note that in C13-pRS and C13-HD6KD cells, equal amounts of immunoprecipitated MSH2, not equal amounts of cell lysates, were loaded on SDS-PAGE.

(D) The HDAC6-selective inhibitor, tubacin, enhances MSH2 acetylation. MEFs were treated with a vehicle control (DMSO), 5 μM tubacin, or 5 mM NaB for 12 hr. IP-WB and direct WB were performed as indicated. For the fourth panel, the original lanes between 1 and 2 were deleted.

(E) SAHA increases the acetylation of MSH2. 293T cells were transfected with HA-MSH2. One sample was treated with a vehicle and the other with 2 μM SAHA overnight. The IP-WB analyses were performed as indicated.

(F) The SirT2 inhibitor, AGK2, does not increase MSH2 acetylation. HeLa cells were transfected with HA-MSH2, then treated with a vehicle control or 25 μM AGK2 (EMD Chemicals) 24 hr prior to harvest. IP-WB and direct WB were performed as indicated.

(G) Depletion of SirT2 in HeLa cells does not increase MSH2 acetylation. HeLa cells were transfected with scramble or SMARTpool siRNA containing four sets of siRNAs against human SirT2 (Dharmacon Inc.) for 3 days. IP-WB and direct WB were performed as indicated. See also Figure S2.

HDAC6 Downregulates MSH2 Protein Stability

In examining the levels of MSH2 in HDAC6WT and HDAC6KO MEFs, we found that MSH2 levels were increased in HDAC6KO MEFs compared with HDAC6WT MEFs (Figure 3A, lanes 1 and 2; Figure 3B, left panel). To determine whether this observation can be made in other cell lines, we examined MSH2 levels in an ovarian cancer cell line pair, C13-pRS (control) and C13-HD6KD, and found that MSH2 protein levels were higher in C13-HD6KD cells compared with the control cells (Figure 3A, lanes 3 and 4; Figure 3B, middle panel). To verify whether HDAC6 could downregulate MSH2 protein levels, wild-type human HDAC6 was introduced into HDAC6KO MEFs. As shown in Figure 3A, lanes 5 and 6, and Figure 3B, in the right panel, HDAC6 indeed reduced MSH2 levels. Therefore, the above data suggest that HDAC6 decreases the protein levels of MSH2.

Figure 3. HDAC6 Downregulates the Protein Levels of MSH2.

Figure 3

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(A) MSH2 levels were detected in three pairs of cell lines: HDAC6WT and HDAC6KO MEFs, C13pRS and C13HD6KD, and empty vector-rescued and F-HDAC6-rescued HDAC6KO MEFs. Direct WB was performed as indicated.

(B) The MSH2 protein levels in (A) were quantified by densitometry. p values were determined by a two-tailed Student’s t test (n = 3).

(C) MSH2’s half-life was examined in five cell lines as indicated. Cell lysates were subjected to anti-MSH2 and anti-β-actin western blotting analyses. The MSH2 bands were quantified by densitometry, and the level of MSH2 at time point zero of HDAC6WT, HDAC6KO, or the rescued clones is designated as 1. The MSH2 level of other time points was quantified, relative to the zero time point, and was shown in fold changes below the panel of anti-MSH2 western blotting analysis.

(D) The MSH6WT and the MSH6KO MEFs were lysed and subjected to western blotting analyses with indicated antibodies.

(E) Each of MSH6KO MEFs and MSH6WT MEFs was transfected with scramble and HDAC6 siRNA. Western blotting analyses were performed as indicated.

(F) The MSH2 protein levels in (E) were quantified as (B).

(G) MSH2’s half-life was detected in two pairs of cell lines as shown in (E). The cells as indicated were either left untreated or treated with 50 μg/ml CHX at the indicated time points. Cell lysates were subjected to anti-MSH2 and anti-β-actin western blotting analyses. The MSH2 bands were quantified by densitometry as in (C). For (B) and (F), data are represented as mean ± SEM. See also Figure S3.

We next examined whether HDAC6-mediated downregulation ofMSH2 is due to decreasing MSH2 protein stability. As shown in Figure 3C, MSH2 exhibited a half-life of 24–36 hr in HDAC6WT MEFs (compare lanes 1, 2, 3, and 4), but one of greater than 36 hr in HDAC6KO MEFs (compare lanes 5, 6, 7, and 8). Therefore, MSH2’s half-life was prolonged in HDAC6KO MEFs, suggesting that HDAC6 plays an important role in destabilizing the MSH2 protein. We then tested whether HDAC6’s deacetylase activity or HDAC6’s DAC1 domain is involved in regulating MSH2 stability. The empty vector, the HDAC6 catalytic dead mutant (F-HD6DM) or the DAC1-deficient mutant (F-HD6 [429–1,215]) was stably transfected into HDAC6KO MEFs, and stable clones expressing the above plasmids were established. As shown in Figure 3C, wild-type HDAC6 (F-HD6, lanes 17–20), not the catalytically dead mutant of HDAC6 (F-HD6DM, lanes 13–16) or the DAC1-deficient mutant of HDAC6 (F-HD6[439–1215], lanes 21–24), could reduce the half-life of MSH2. This indicates that either HDAC6’s deacetylase activity or HDAC6’s DAC1 domain is necessary for controlling MSH2 protein stability in vivo. In addition, the ZnF-UBP domain of HDAC6 appears to be important in affecting MSH2’s half-life (Figure S3A).

Previous studies suggest that MSH6 stabilizes MSH2 by forming MSH2-MSH6 heterodimers (Hayes et al., 2009). Consistently, the levels of MSH2 are dramatically decreased in MSH6KO MEFs compared with their counterparts (Figure 3D). Therefore, we assumed that there was more monomeric MSH2 in MSH6KO MEFs than that in MSH6WT MEFs. We then hypothesized that HDAC6 destabilizes MSH2 more pronouncedly in MSH6KO than in MSH6WT MEFs. Supporting this hypothesis, as shown in Figures 3E and 3F, transient knockdown of HDAC6 in MSH6KO MEFs caused a more than 5-fold increase in MSH2 compared with that in control cells. In contrast, transient knockdown of HDAC6 in MSH6WT MEFs only caused a less than 2-fold MSH2 increase compared with the control. As expected, in MSH6KO MEFs MSH2’s half-life (less than 2 hr) is much shorter than that (greater than 8 hr) in MSH6WT MEFs (Figure 3G, lanes 1–5 versus lanes 11–15). Moreover, knockdown of HDAC6 in MSH6KO MEFs dramatically prolonged MSH2’s half-life (Figure 3G, lanes 6–10 versus lanes 1–5) compared with that in MSH6WT MEFs (Figure 3G, lanes 16–20 versus lanes 11–15). Therefore, these results indicate that HDAC6 markedly promotes monomeric MSH2 degradation.

HDAC6’s DAC1 Domain Functions as an E3 Ligase to Ubiquitinate MSH2 In Vitro

It has been documented that MSH2 is degraded via the ubiquitin- proteasome pathway (Hernandez-Pigeon et al., 2004). Therefore, we investigated whether HDAC6 modulates MSH2 stability through this pathway. As shown in Figure 4A, MSH2 accumulated in HDAC6WT MEFs treated with the proteasome inhibitor MG132 (lanes 1 and 2), but not in the treated HDAC6KO MEFs (lanes 3 and 4), suggesting that MSH2’s turnover is regulated by the ubiquitin-proteasome pathway in an HDAC6-dependent manner. We then tested whether HDAC6 is essential for MSH2 ubiquitination. MSH2 from HDAC6WT or HDAC6KO MEFs was immunoprecipitated by anti-MSH2 antibodies, and polyubiquitination was detected by anti-ubiquitin antibodies. As shown in Figure 4B, upon MG132 treatment, MSH2 polyubiquitination was abolished in HDAC6KO MEFs (lane 4) as compared with that in HDAC6WT MEFs (lane 2). Furthermore, MSH2 polyubiquitination was restored by reintroduction of wild-type HDAC6 (lane 10), not the catalytically dead mutant of HDAC6 (lane 8). These data indicate that HDAC6’s deacetylase activity is critical for MSH2 polyubiquitination in vivo.

Figure 4. HDAC6 Is an Ubiquitin E3 Ligase Targeting MSH2.

Figure 4

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(A) HDAC6WT and HDAC6KO MEFs were left untreated or treated with 10 μM MG132 for 12 hr. Anti-MSH2 and anti-β-actin western blotting analyses were carried out.

(B) MSH2 ubiquitination correlates with the presence of wild-type HDAC6. Five cell lines, HDAC6WT MEFs, HDAC6KO MEFs, and empty vector-, F-HD6DM-, and F-HD6-rescued HDAC6KO MEFs, were left either untreated or treated with 20 μM MG132 for 4 hr. IP-WB was performed as indicated. The original lanes between lanes 4 and 5 and lanes 6 and 7 were deleted.

(C) HDAC6 is autoubiquitinated. Ubiquitination assays were set up with all the components: ATP, HA-Ub, E1, E2 (UbcH5a), and insect cell-expressed GST-HDAC6 (lane 6) as described in the Supplemental Experimental Procedures. Anti-HA western blotting analysis was performed.

(D) HDAC6 ubiquitinates MSH2 in vitro. Ubiquitination assays were performed as described in the Supplemental Experimental Procedures. The Ni-NTA bead-bound MSH2 was washed under the denaturing condition as described in the Supplemental Experimental Procedures. Anti-HA western blotting analysis was performed.

(E) Gel-purified HDAC6 harbors an E3 ligase activity. Boiled or renatured F-HDAC6 was prepared as described in the Supplemental Experimental Procedures. The autoubiquitination assay was then performed, and the polyubiquitinated HDAC6 was detected by western blotting analysis using anti-HA antibodies.

(F) In vitro ubiquitination assays were set up as described in (C), except that HA-Ub was replaced by His-Ub, (lanes 1 and 3), His-K48R (lane 4), or His-K63R (lane 5) in the reaction. Anti-His western blotting analysis was carried out.

(G) The in vitro ubiquitination assays were set up as (F) using Flag-HDAC6 purified from Sf9 cells as an E3 enzyme and the bacterially purified GST-MSH2 as a substrate. An anti-His western blotting analysis was performed to determine the polyubiquitination of GST-MSH2.

(H) Diagrams of Flag-tagged DAC1 and DAC2 deletion proteins expressed in Sf9 cells.

(I) The ubiquitination reactions were carried out as in (C) except that F-DAC1 or F-DAC2 purified from Sf9 cells was used to replace GST-HDAC6, and GST-MSH2 was used to replace His-MSH2. The ubiquitination status of GST-MSH2 was detected by anti-HA antibodies. The protein levels of DAC1 and DAC2 in Ub assays and HDAC assays were indicated by Coomassie blue staining (lower panel). The original lanes between lanes M and 1 and lanes 1 and 2 were deleted.

(J) HDAC enzymatic activities of DAC1 and DAC2 were determined by HDAC assays performed using 3[H]-labeled core histones as substrates as described in the Experimental Procedures. The data are represented as mean ± SEM.

(K) Ubiquitination assays were set up as in (D), except that F-HD6(1–503) was used to replace GST-HDAC6, and GST-MSH2 replaced His-MSH2. Ubiquitinated MSH2 was examined by an anti-HA western blotting analysis. See also Figure S3.

Based on the above data, we hypothesize that HDAC6 ubiquitinates MSH2 as an E3 ligase. Supporting our hypothesis, insect cell-expressed recombinant GST-HDAC6 is efficiently autoubiquitinated, which is a hallmark of E3 ligases. As shown in Figure 4C, in the presence of ATP, Ub, E1, and E2 (UbcH5a), HDAC6 itself can be ubiquitinated (lane 6), suggesting that the ubiqutination signal is indeed from HDAC6. To determine which E2 is recruited for the ubiquitination of MSH2, we screened a panel of E2s and found that HDAC6’s E3 ligase activities depend on the UbcH5 subfamily (Figure S3B). For simplicity, UbcH5a was chosen to be used in the later experiments. We next examined whether HDAC6 serves as an E3 ligase for MSH2. In the presence of ATP, Ub, E1, E2, and insect cell-expressed GST-HDAC6, bacterially purified His-MSH2 was polyubiquitinated under denaturing conditions (Figure 4D, lane 7), which confirmed MSH2 as the source of the ubiquitination signal. To exclude the possibility of contaminating E3 ligase activity, F-HDAC6 overexpressed in 293T cells was gel purified, extracted, and then renatured. As shown in Figure 4E, renatured but not boiled HDAC6 was self-ubiquitinated in the reaction, indicating that HDAC6 itself harbors an intrinsic E3 ligase activity and that this activity is conformation dependent. To further determine whether the ubquitinated ladders are from polyubiquitination or monoubiquitination, we used the recombinant ubiquitin mutant K48R or K63R to replace the wild-type ubiquitin in the reaction. As shown in Figure 4F, the K48R mutant completely abolished the polyubiquitination of GST-HDAC6 purified from Sf9 cells (lane 4), whereas K63R had no effect (lane 5), indicating that HDAC6 promotes polyubiquitination specifically through the formation of the K48 linkage. Consistently, HDAC6 also promotes MSH2 polyubiquitination through the formation of the K48 linkage (Figure 4G).

Since HDAC6 decreases MSH2’s half-life more pronouncedly in MSH6KO than in MSH6WT MEFs, we speculated that HDAC6 would promote ubiquitination more efficiently in monomeric MSH2 than in the MutSα and MutSβ complexes. To test our hypothesis, we performed in vitro E3 assays using Sf9 purified His-MSH2, MutSα, and MutSβ as substrates. As shown in Figure S3C, HDAC6 promoted robust ubiquitination in monomeric MSH2, but only very weak ubiquitination in MutSα and MutSβ, suggesting that HDAC6 prefers to ubiquitinate monomeric MSH2 rather than MutSα and MutSβ. We then examined which region of HDAC6 harbors an E3 ligase activity toward nonacetylated MSH2. The insect cell-purified DAC1 or DAC2 domain was used in the in vitro ubiquitination reactions (Figure 4H). DAC1 efficiently polyubiquitinated MSH2 (Figure 4I, lane 2); in contrast, DAC2 did not (Figure 4I, lane 3). Conversely, DAC2 displayed strong deacetylase activity, while DAC1 was catalytically dead (Figure 4J). In agreement with this, DAC1 purified from 293T cells possessed ubiquitination activity toward GST-MSH2 in vitro (Figure 4K, lane 7). These data indicate that the DAC1 domain harbors an intrinsic E3 ligase activity.

HDAC6 Sequentially Deacetylates and Ubiquitinates MSH2 In Vivo

To determine whether HDAC6 promotes MSH2 ubiquitination in vivo, 293T cells were cotransfected with His-MSH2 combined with wild-type HDAC6 (F-HD6), a DAC2 catalytic site mutant of HDAC6 (F-HD6[H611A]) or DAC1-deficient HDAC6 (F-HD6 [439–1,215]). As shown in Figure 5A, F-HD6, but not F-HD6(H611A) or F-HD6(439-1215), is able to increase the ubiquitination signal of His-MSH2 in vivo. In addition, the F-HD6 (439-1215)-rescued clone could not reduce MSH2’s half-life in HD6KO MEFs (Figure 3C, lanes 21–24), suggesting that both HDAC6’s deacetylase activity and DAC1’s E3 ligase activity are essential for MSH2 ubiquitination and degradation in vivo.

Figure 5. HDAC6 Sequentially Deacetylates and Ubiquitinates MSH2.

Figure 5

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(A) 293T cells were cotransfected with His-MSH2 and either an empty vector, F-HD6, F-HD6(H611A), or F-HD6(439–1215). Ni-NTA beads were used to pull down His-MSH2 as described in the Supplemental Experimental Procedures. Western blotting analyses were performed as indicated.

(B) For lanes 1–4, ubiquitination assays were set up with rabbit reticulocyte lysate (RRL) (a source of E1, E2, and ATP) and GST-HDAC6 purified from Sf9 cells (lane 1), RRL, Flag-Ub and GST (lane 2), RRL, Flag-Ub, and GST-HDAC6 (lane 3) or RRL, Flag-Ub, GST-HDAC6 with TSA (2 μg/ml) (lane 4). IPed MSH2 from HDAC6KO MEFs served as the substrate. The ubiquitination status of MSH2 was examined by anti-Flag western blotting analysis. For lanes 5–7, in vitro ubiquitination reactions were set up as in the left panel except that bacterially purified GST-MSH2 served as the substrate and HA-Ub was used to replace Flag-Ub. Sf9-purified HDAC6 was absent (lane 5), or present (lane 6), or present with 2 μg/ml TSA (lane 7). The ubiquitination status of MSH2 was examined by anti-HA western blotting analysis.

(C) GST-HDAC6 employed in (B) was tested for its ability to deacetylate core histones in the absence or presence of 2 μg/ml TSA in standard HDAC assays as described in the Experimental Procedures.

(D) Diagrams of Sf9 cell-purified F-HD6 and F-HD6DM proteins.

(E) Ubiquitination assays were set up as in (B). HDAC6 deacetylase catalytic dead mutant can ubiquitinate MSH2 purified from bacteria, but not from HDAC6KO MEFs. For lanes 1 and 4, reaction buffer (ctl) was used to replace the recombinant HDAC6. The substrates of the ubiquitination assays are indicated above the gel. The ubiquitination status of MSH2 was examined by an anti-HA western blotting analysis.

(F) The protein levels of HDAC6 WT and HDAC6 DM are shown by Coomassie blue staining. The original lanes between the marker and the F-HD6 were deleted.

(G) The HDAC assays were performed as described in Experimental Procedures. For (C) and (G), data are presented as mean ± SEM.

To determine how HDAC6’s deacetylase activity and its E3 ligase activity work in concert to ubiquitinate MSH2 in vivo, we used a pan-HDAC inhibitor, trichostatin A (TSA), and a deacetylase dead mutant to examine the impact of HDAC6’s deacetylase activities on the ubiqutination of MSH2. As shown in Figures 5B and 5C, the ability of GST-HDAC6 purified from Sf9 cells to ubiquitinate MSH2 isolated from HDAC6KO cells was markedly reduced in the presence of TSA (lane 4), which inhibits HDAC6’s deacetylase activity, compared with the control (lane 3). Conversely, GST-HDAC6 was able to efficiently ubiquitinate MSH2 isolated from E. coli in the presence of TSA (lane 7). These results indicate that HDAC6’s E3 ligase activities toward nonacetylated MSH2 are independent of its deacetylase activity. To corroborate our results, we employed the wild-type and the catalytically dead mutant of HDAC6 purified from Sf9 cells to perform the in vitro ubiquitination assays. As shown in Figures 5D–5G, wild-type but not catalytically dead mutant of HDAC6 could efficiently ubiquitinate MSH2 isolated from KO MEFs containing acetylated MSH2 existing as MutSα (Figure 5E, lanes 2 and 3). In contrast, both wild-type and the catalytically dead mutant of HDAC6 could promote polyubiquitination of MSH2 isolated from bacteria existing as the nonacetylated MSH2 monomers (lanes 5 and 6). These data strongly suggest that HDAC6 sequentially deacetylates and polyubiquitinates MSH2 in vivo.

Lysines 845, 847, 871, and 892 ofMSH2Are Targeted for Acetylation as well as Ubiquitination

Protein acetylation often influences protein stability (Sadoul et al., 2008). To test whether protein acetylation affects MSH2 stability, we examined MSH2’s half-life and acetylated MSH2’s half-life. HA-MSH2 was overexpressed in 293T cells followed by cycloheximide (CHX) treatment. As shown in Figure 6A, in the top panel, the half-life of the total MSH2 is approximately 1 hr. Acetylated MSH2 was examined by IP with anti-HA antibodies and western blotting with anti-AcK antibodies. As shown in Figure 6A, in the middle panel, the half-life of acetylated MSH2 is longer than 4 hr. Since most of MSH2 in cells is heterodimerized with MSH6 or MSH3, we assume that the overexpressed MSH2 exists as monomers. The half-life we measured in Figure 6A is the half-life of monomeric MSH2. Therefore, these data suggest that acetylation of monomeric MSH2 stabilizes the protein. We then set out to identify the acetylation sites in MSH2 by overexpression of MSH2 in 293T cells followed by treatment with TSA and mass spectrometry analyses. As shown in Figures S4 and 6B, K845, K847, K871, and K892 were detected as acetylation sites. To ensure that these four sites are indeed in vivo acetylation sites, we mutated them into arginine (4KR). As shown in Figure 6C, the level of 4KR acetylation was decreased as compared to wild-type, suggesting that these sites were acetylated in vivo. Since MSH2(4KR) is still acetylated, we predict that there are yet-unidentified acetylation sites in MSH2. Because HDAC6 also targets MSH2 for ubiquitination, we overexpressed HA-MSH2 in 293T cells followed by MG132 treatment, and then subjected them to mass spectrometry analyses. We found that the above acetylation sites could also be ubiquitinated when proteasome degradation was inhibited (Figures S5 and 6B). We then investigated whether these four sites are ubiquitinated in vivo. As shown in Figure 6D, the 4KR mutant exhibited reduced ubiquitination compared with the wild-type (lane 4 versus lane 3), suggesting that these four lysine sites are ubiquitinated in vivo.

Figure 6. Lysines 845, 847, 871, and 892 of MSH2 Are Targeted for Acetylation as well as Ubiquitination.

Figure 6

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(A) The half-life of acetylated MSH2 is longer than that of total MSH2. 293T cells were transfected with HA-MSH2. After 36 hr, cells were treated with cycloheximide (CHX) as indicated. IP-WB and WB were performed as indicated. The bands were quantified by densitometry as in Figure 3C.

(B) The diagram of MSH2 showing MSH2 domains and overlapped acetylated/ubiquitination sites.

(C) The four lysine sites are acetylated in vivo. 293T cells were transfected with HA-MSH2 or HA-MSH2(4KR). Cells were treated with TSA for 4 hr prior to harvest. HA-MSH2 and HA-MSH2(4KR) were IPed by anti-HA agarose beads followed by an anti-AcK western blotting analysis. The expression levels of HA-MSH2 and HA-MSH2(4KR) were detected by Ponceau staining.

(D) The four lysine sites are ubiquitinated in vivo. His-Flag-Ub was cotransfected with either HA-MSH2-C(WT) or HA-MSH2-C(4KR) into 293T cells. The transfectants were treated with either a vehicle or MG132 for 4 hr prior to harvest. Ni-NTA beads were used to pull down His-Flag-Ub. An anti-Flag western blotting analysis was performed. Arrows indicate the nonspecific bands.

(E) For lanes 1–11 and 17–20, 293T cells were cotransfected with the indicated plasmids. Anti-HA agarose beads were used to immunoprecipitate the wild-type or mutants of HA-MSH2. Anti-Flag and anti-HA western blotting analyses were performed using the anti-HA immunoprecipitates. Anti-HA and anti-Flag western blotting analyses were also performed using the cell lysates. For lanes 12–16, LoVo cells were transfected with the indicated plasmids. IP-WB and WB experiments were performed as indicated in 293T cells.

(F) The MSH2(4KR) mutant reduces the interaction with HDAC6 compared with the MSH2 wild-type. 293T cells were transfected with the indicated plasmids. IP-WB and WB were performed as indicated.

(G) 293T cells were transfected with either HA-MSH2(WT) or HA-MSH2(4KR) followed by CHX treatment at indicated time intervals. Anti-HA and anti-β-actin western blotting analyses were performed. The HA-MSH2 bands were quantified by densitometry as in Figure 3C. See also Figures S3–S5 and S7.

We next examined the possible functions of the acetylation/ubiquitination of these four lysine sites. Previous studies showed that the C-terminal region of MSH2 is important for MSH6 and MSH3 binding (Guerrette et al., 1998). We then examined whether the 4KR mutant, a deacetylation-mimicking mutant, affects the MSH2-MSH6 interaction. As shown in Figure 6E, the MSH2(4KR) mutant abolished its binding to MSH6 (lane 3), but not to MSH3 (lane 19), compared with the wild-type (lanes 2 and 18), suggesting that deacetylation of these four lysine sites may disrupt MutSα, but not MutSβ, complexes. We next examined which lysine is important for MutSα assembly. So the above four lysines were substituted to arginine individually (K845R, K847R, K871R, and K892R) or in combination (K845/847R). As shown in Figure 6E, K845R, K847R, and K845/K847R slightly reduced their binding to MSH6 compared to the wild-type (compare lanes 5, 6, 8, and 10), while K871R and K892R did not affect their binding to MSH6 compared to the wild-type (compare lanes 12, 13, and 15). Intriguingly, MSH2(4KQ) (an acetylation-mimicking mutant, where the lysine is substituted to a glutamine), also disrupted its binding to MSH6 (lanes 2 and 4), but not to MSH3 (lanes 18 and 20). Moreover, similar to the above K to R mutants, the mutants of K845Q, K847Q, and K845/K847Q slightly reduced their binding to MSH6 compared with the wild-type (compare lanes 5, 7, 9, and 11), while K871Q and K892Q did not affect their binding to MSH6 compared to the wild-type (compare lanes 12, 14, and 16). Thus, there are two possibilities which can account for these observations. First, the 4KQ mutant cannot faithfully mimic the acetylation status. Second, these four lysine sites per se, not their modifications (acetylation/deacetylation), are important for MutSα assembly. Recently, a computational model showed that MSH2 with an acetylated K845 has more favorable dimerization energy (Figure S3D), suggesting that acetylation of MSH2 facilitates the MutSα complex assembly. This model favors the first possibility. Our recent results showed that purified HDAC6 can disrupt the MutSα complex and that depletion of HDAC6, or inhibition of HDAC6 activity, causes an increased MutSα complex formation (Figure S7A), suggesting that deacetylation of MSH2 by HDAC6 facilitates the disassembly of MutSα. Additionally, even if further work shows that the second possibility is correct, there is a likelihood that MutSα disassociation could also be due to deacetylation of yet-unidentified lysines in MSH2. We also examined whether MSH2(4KR) (the deacetylation- mimicking mutant) affects its interaction with HDAC6. As shown in Figure 6F, this mutant displays a reduced interaction with HDAC6 compared with the wild-type, suggesting that HDAC6 may interact with the acetylated form of MSH2 stronger than the deacetylated form of MSH2. We next tested whether MSH2(4KR), which can also be referred to as a nonubiquitinated mutant, would prolong the half-life of MSH2. As shown in Figure 6G, the half-life of MSH2(4KR) is much longer than that of the MSH2(WT) (8 hr versus 4 hr). Thus, ubiquitination of these four lysine sites plays an important role in destabilizing the MSH2 protein.

HDAC6 Causes a Cellular Tolerance to 6-TG and MNNG and Decreased Cellular MMR Activity by Downregulation of MSH2

It has been well documented that the levels of MSH2 are inversely correlated with 6-TG and MNNG resistance (Karran, 2006). Consistent with this observation, HDAC6KO MEFs harboring more MSH2 were more sensitive to 6-TG and MNNG by MTT assays (Figures S6A and S6D). We then investigated whether the reduced cell numbers in the presence of 6-TG and MNNG were due to apoptosis using poly ADP-ribose polymerase 1 (PARP1) cleavage and caspase-3 cleavage as readouts. HDAC6KO MEFs and HDAC6KD C13 exhibited higher sensitivity to 6-TG- and MNNG-induced apoptosis compared with control cells (Figures S6B, S6C, and S6E). To confirm that HDAC6 is associated with a long-term 6-TG resistance, colony-formation assays were carried out. As shown in Figure S6F, there was less colony formation in HDAC6KO MEFs upon 6-TG treatment compared with that in HDAC6WT MEFs, while MSH2KO MEFs exhibited more colonies after 6-TG treatment as compared with MSH2WT MEFs (Figure S6G). These results indicate that the loss of HDAC6 sensitizes cells to 6-TG via increasing the levels of MSH2. We next determined whether HDAC6’s deacetylase activities are essential for regulating 6-TG sensitivity. As shown in Figure 7A, the HDAC6 wild-type-rescued (lanes 10, 11, and 12), not the HDAC6 dead mutant-rescued HDAC6KO MEFs (lanes 7, 8, and 9), displayed diminished sensitivity to 6-TG compared to the empty vector-rescued HDAC6KO MEFs, suggesting that HDAC6’s deacetylase activities are essential for governing 6-TG sensitivity. We next investigated whether HDAC6 mediates 6-TG resistance via MSH2. To this end, HDAC6KOMEFs were depleted ofMSH2by shRNA (Figure S7B). As shown in Figure 7B, depletion of MSH2 in an HDAC6 null background led to delayed apoptosis as indicated by the PARP1 cleavage bands compared with the control cells (lanes 11 and 12 versus lanes 7 and 8). The above data suggest that HDAC6 functions upstream of MSH2. To determine whether the levels of MutSα in cells are important for cellular 6-TG sensitivity, F-MSH6 was cotransfected with wild-type MSH2 or MSH2(4KR) into MSH2-deficient LoVo cells, which did not express detectable levels of MSH6 either. As shown in Figure 7C, cells transfected with 4KR displayed more tolerance to 6-TG treatment. Since the MSH2(4KR) mutant lacks the ability to form the MutSα complex (Figure 6E), this observation suggests that low levels of MutSα lead to cellular tolerance to 6-TG. To confirm the above results, MSH2(WT)- and MSH2(4KR)-rescued MSH2KO MEF pools were established and colony-formation assays were performed (Figures 7D–7F). As expected, MSH2(4KR)-rescued MEFs were more resistant to 6-TG compared with the MSH2(WT)-rescued MEFs (Figures 7E and 7F), suggesting that low levels of MutSα are associated with cellular tolerance to DNA damage.

Figure 7. HDAC6 Confers an Increased Cellular Resistance to 6-TG and Decreased DNA Mismatch Activity.

Figure 7

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(A) HDAC6 deacetylase activity is required to regulate cellular sensitivity to 6-TG. HDAC6WT MEFs and vector-, F-HD6DM-, and F-HD6-rescued HDAC6KO MEFs were treated with indicated 6-TG for different time intervals. Anti-PARP1 and anti-β-actin western blotting analyses were performed. The original lanes between lanes 3 and 4 and lanes 9 and 10 were deleted.

(B) MSH2 functions as a downstream target of HDAC6. HDAC6WT and HDAC6KO MEF clones harboring scramble shRNA or MSH2 shRNA (Santa Cruz, CA) were treated with 30 μM 6-TG at indicated time intervals. Anti-PARP1 and anti-β-actin western blotting analyses were carried out. The original lanes between lanes 4 and 5 and lanes 8 and 9 were deleted in the upper panel.

(C) MSH2 (4KR) confers cellular resistance to 6-TG. F-MSH6 was transiently transfected along with either HA-MSH2(WT) or HA-MSH2(4KR) into LoVo cells followed by 6-TG (60 μM) treatment as indicated by time intervals. Western blotting analyses were performed as indicated.

(D) Anti-HA and anti-β-actin western blotting analyses were performed using the lysates from HA-MSH2(WT)- and HA-MSH2(4KR)-rescued MSH2KO MEFs.

(E) A representative snapshot of the colony-formation assay.

(F) Colony-formation assays of HA-MSH2(WT)- and HA-MSH2(4KR)-rescued MSH2KO MEFs were performed as described in the Supplemental Experimental Procedures. Statistical analysis was performed using the Student’s test (p = 0.00017). A p value less than 0.05 was considered significant. An asterisk (*) represents the p value. Data are presented as mean ± SEM

(G) (Upper panel) 293T cells were cotransfected with His-MSH2 and HA-Ub followed by 6-TG treatment as indicated. IP-WB and direct WB were performed as indicated. (Lower panel) 293T cells were treated by 6-TG as indicated. Western blotting analyses were performed with indicated antibodies.

(H) 293T cells were cotransfected with HA-MSH2 and Flag-MSH6 followed by 6-TG treatment at indicated time intervals. IP-WB and direct WB were performed as indicated.

(I) The principle of the in vitro MMR assay. Please see the Supplemental Experimental Procedures for details.

(J) (Left panel) HeLa (as a positive control), HDAC6WT, and HDAC6KO MEF NEs were incubated with the above DNA substrate followed by digestion with HindIII and BspD1. Unrepaired and repaired products were indicated and quantified by densitometry. (Right panel) An independent replicate. See also Figures S6 and S7.

We next examined whether MSH2’s deacetylation and ubiquitination by HDAC6 are modulated by DNA damage. As shown in Figures 7G and S7C, upon 6-TG treatment, ubiquitination of MSH2 was reduced and acetylation of MSH2 at K847 was increased, implying that the levels of MSH2 are increased. Interestingly, the acetylation level of tubulin was increased after 6-TG treatment, suggesting that HDAC6’s deacetylase activity is decreased upon DNA damage. In addition, as shown in Figure 7H, following 6-TG treatment the levels of MutSα were increased. The above results suggest the model that DNA damage may decrease HDAC6’s deacetylase and E3 ligase activities, facilitate MutSα assembly, and lead to apoptosis in cells. Besides HDAC6’s role in DNA damage response, its role in DNA mismatch repair was also explored. As shown in Figures 7I and 7J, HDAC6KO MEFs harbored higher MMR activities compared with HDAC6WT MEFs, which is consistent with the data showing that higher levels of MutSα exist in HDAC6KO MEFs compared with that in HDAC6WT MEFs (Figures 3A and S7A, lane 4 versus lane 3). Since the 4KR mutant does not bind to MSH6 (Figure 6E), and since MutSα (MSH2-MSH6) only functions as a heterodimer, not an MSH2 or MSH6 monomer (Drummond et al., 1995), we expect that the 4KR mutant in MSH2KO MEFs will not execute MMR function. Consistent with our hypothesis, the 4KR-rescued MSH2KO MEFs exhibited more colonies upon 6-TG treatment compared with the WT-rescued MSH2KO MEFs in a colony-formation assay (Figures 7D–7F), suggesting that the 4KR-rescued MSH2KO MEFs harbor much lower MMR activities compared with the WT-rescued ones. In conclusion, HDAC6 plays an important role in both DNA damage response and DNA mismatch repair by regulating the levels of MutSα.

DISCUSSION

We and others have recently shown that absence or depletion of HDAC6 sensitizes cells to DNA-damaging anticancer agents, such as cisplatin and doxorubicin (Namdar et al., 2010; Wang et al., 2012). However, the mechanisms behind this observation are unclear. In this paper, we report that HDAC6-induced cellular tolerance to 6-TG and MNNG is partly attributed to HDAC6-mediated downregulation of the MutSα complex. Briefly, our studies support a model in which HDAC6 first targets MutSα to deacetylate MSH2, causing MSH2 disassociation from MSH6, a binding partner that stabilizes MSH2 (Chang et al., 2000; Hayes et al., 2009). HDAC6 then targets the resulting monomeric MSH2 to promote its ubiquitination and degradation, which ultimately leads to the reduced levels of MutSα and cellular tolerance to DNA-damaging agents. However, we cannot rule out the possibility that HDAC6 may also directly deacetylate and ubiquitinate monomeric MSH2, leading to a decrease in the levels of MutSα. These two models are not mutually exclusive.

Under static conditions, the levels of nuclear HDAC6 protein are generally lower than the levels of cytoplasmic HDAC6 (Verdel et al., 2000). This may be partly attributed to the SE14 domain of human HDAC6, which serves as a cytoplasmic anchor of the protein (Bertos et al., 2004). However, mouse HDAC6 has been reported to enter the nucleus under specific circumstances, such as after butyrate-induced B16 cell differentiation (Verdel et al., 2000). We have also found that upon treatment with UV or DNA-damaging agents, a fraction of HDAC6 translocates into the nucleus (Figure S7D). Interestingly, in response to alkylating agents, MutSα can also be accumulated in the nucleus (Christmann and Kaina, 2000). It is yet undetermined whether HDAC6 plays a role in MutSα translocation.

Among 18 deacetylases, HDAC6 is the one most intimately associated with ubiquitin. Structurally, HDAC6 harbors a unique Zn-UBP domain which binds to monomeric ubiquitin with the highest known affinity for ubiquitin binding among all known ubiquitin-interacting proteins (Boyault et al., 2006). The Zn-UBP domain also binds to ubiquitinated proteins. In particular, the Zn-UBP domain interacts with misfolded and polyubiquitinated proteins (Kawaguchi et al., 2003). A small region between the DAC1 and DAC2 domain of HDAC6 has been reported to simultaneously bind to dynein motors and transport the misfolded proteins to aggresomes for autophagy, a lysosome-mediated degradation pathway (Kawaguchi et al., 2003). Therefore, HDAC6 is believed to play a protective role in response to the accumulation of cytotoxic protein aggregates. Additionally, in a Drosophila model, autophagy is compensated for by impairment of the ubiquitin-proteasome system (UPS) in an HDAC6-dependent manner (Pandey et al., 2007). Emerging evidence suggests that HDAC6 may also be involved in an ubiquitin-proteasome pathway. A recent investigation has shown that HDAC6 promotes polyubiquitination of Cdc20, a coactivator for the anaphase-promoting complex (APC), and enhances the activity of the Cdc20-APC complex (Kim et al., 2009). This report has suggested that HDAC6 may be one of the components in the ubiquitin-proteasome pathway that ubiquitinates Cdc20, and that HDAC6’s Zn-UBP domain, not its deacetylase activity, is required for HDAC6-mediated Cdc20 ubiquitination. However, no rigorous experiments were performed to demonstrate that HDAC6 functions as an E3 ligase. In the present study, we have demonstrated that recombinant HDAC6 exhibits intrinsic ubiquitin E3 ligase activity toward MSH2. The HDAC6 N-terminal DAC1 domain (aa 1–503) is sufficient to ubiquitinate unacetylated MSH2 in vitro, but HDAC6’s deacetylase activity and Zn-UBP domain may be required to ubiquitinate acetylated MSH2 in vivo (Figures 4 and S3A). Future study will be focused on the delineation of the key amino acids which are important for the E3 ligase activity in the N terminus of HDAC6. A complete understanding of the structure of the HDAC6’s DAC1 domain will be needed to achieve this goal. Therefore, HDAC6 appears to be a vital player in both protein degradation pathways: the autophagy and the ubiquitin-proteasome pathways.

As a key DNA mismatch repair protein, little is known about MSH2’s posttranslational modification. Our study is the first to show that MSH2 is acetylated at four lysine residues (K845, K847, K871, and K892). Interestingly, these four sites could also be ubiquitinated. A deacetylation mimetic mutant of MSH2 (4KR) dramatically disrupts the MutSα complex (Figure 6E). Intriguingly, an acetylation mimetic mutant (4KQ) disrupts the MutSα complex as well (Figure 6E). It is possible that this acetylation mimetic mutant may not faithfully mimic the acetylation status. Previously, Guerrette et al. (1998) mapped the MSH6 binding region to the 70 aa (875–934) within the C terminus of MSH2 (Guerrette et al., 1998). Among the four acetylated lysines, K892 was the only one within this region. However, the single mutations K892R and K892Q did not affect the MSH2-MSH6 association (Figure 6E). One of the acetylation/ubiquitination sites, K845, has been found to be a missense mutation in hereditary nonpolyposis colorectal cancer (HNPCC) (Nomura et al., 2000). No change in the interaction between this mutation (K845E) and MSH6 was found in yeast (Gammie et al., 2007). However, K845E exhibited decreased DNA MMR activities compared with the wild-type in yeast (Gammie et al., 2007), suggesting that acetylation/ubiquitination of MSH2 at K845 may affect its MMR activity. In addition, computational modeling has shown that the computed dimerization energies are overwhelmingly in favor of acetylated MSH2(K845) to form MutSα (Figure S3D). Consistently, K845R slightly reduced their interaction with MSH6 compared with WT (Figure 6E, lanes 5 and 6). Nevertheless, our results suggest that all four lysine sites are needed to be deacetylated to promote disassembly of MutSα. Besides the four acetylation sites shown here, additional sites were also identified by mass spectrometry. Elucidating the role of those additional acetylation sites will be of great significance.

EXPERIMENTAL PROCEDURES

Purification of HDAC6 Nuclear Complex

For Figure 1A, six grams of HeLaS3 NE were loaded onto a 700 ml P11 column equilibrated with buffer C (20 mM Tris-HCl [pH 7.9], 0.2 mM EDTA, 1 mM DTT, 0.1mMPMSF, 0.025% NP-40, 10% glycerol) containing 100mMKCl (BC100). Proteins that bound to the column were step eluted with BC300, BC500, and BC1000. The BC300 fraction, containing HDAC6, was dialyzed against Buffer D (40 mM HEPES-KOH [pH 7.9], 0.2 mM EDTA, 1 mM DTT, 0.1 mM PMSF, 0.025% NP-40, 10% glycerol) containing 20 mM ammonium sulfate (BD20) and then loaded onto a 45 ml HPLC-DEAE-5PW column (TOSOH Bioscience). Bound proteins were eluted with 8 column volume (cv) linear gradient from BD20 to BD500. Presence of HDAC6 in the fractions was detected by the western blot analysis. HDAC6 was observed in the 133–270mMfraction which was then subsequently loaded onto a Superose 6 column, or affinity purified by anti-HDAC6 antibodies.

Immunoprecipitation and Immunoblotting

For immunoprecipitations, cells were lysed in LS buffer (PBS [pH 7.5], 10% glycerol, 0.1% NP-40, PMSF, and protease inhibitor cocktail). Lysates were incubated with protein A or protein G agarose (Invitrogen) for 2 hr for preclearing prior to incubation with the indicated primary antibodies for 12 hr at 4°C. Immunocomplexes were collected, washed four times in TBST buffer (0.1% Tween-20 in TBS), and resolved by SDS-PAGE. For immunoblotting, samples were transferred to nitrocellulose membranes that were then probed with the indicated antibodies. Bound antibodies were detected using a Chemiluminescent Detection Kit (Pierce).

In Vivo Ubiquitination Assay

For Figure 5A, His-MSH2 (2 μg) was cotransfected with 4 μg of vector, F-HD6, F-HD6(H611A), or F-HD6(439-1215) into 293T cells. Thirty-six hours posttransfection, cells were harvested. Cell pellets were lysed in Buffer A, 30 μl Ni-NTA agarose beads were added, and the mixture was rotated at RT for 12 hr. The beads were sequentially washed and resolved on SDS-PAGE as in the above in vitro ubiquitination assays.

Histone Deacetylation Assay

For Figures 4J, 5C, and 5G, the HDAC6 full-length, HDAC6 dead mutant, and HDAC6 deletion proteins purified from Sf9 cells were assayed for their deacetylase activities as described in Tsai and Seto (2002). Briefly, [3H]acetate-incorporated histones were isolated from butyrate-treated HeLa cells by acid extraction as described (Carmen et al., 1996). Purified core histones (12,000 cpm) were incubated with Sf9 cell-purified proteins in 150 μl of ice-cold HD buffer (20 mM Tris [pH 8.0], 150 mM NaCl, and 10% glycerol) at 30°C for 2hr. The reaction was terminated by the addition of an equal volume of stop solution (0.16 M acetic acid, 1.0 M HCl) and mixed well by vortexing. The released [3H]acetate was extracted with ethyl acetate and combined with a scintillation mixture for analysis.

Supplementary Material

Sup1
Sup2

Acknowledgments

We thank Drs. Saadi Khochbin, Yi Zhang, and Nancy Olashaw for discussion, critical reading, and editing of this manuscript. We thank Drs. Jia Fang, X.-J. Yang, Meera Nanjundan, T.-P. Yao, Stuart L. Schreiber, Ralph Mazitschek, James E. Bradner, and Frank Jirik for reagents, and ICG for funding. We also thank Ms. Wei Guan and Ms. Victoria Izumi at Moffitt Cancer Center for technical assistance. H.W. is a Leukemia and Lymphoma Scholar and is supported by NIH grant GM081489. X.Z. is a K30 Scholar and a Liz Tilberis Scholar. K.A.W. is supported by a USF Graduate Student Success Diversity Fellowship. This work is supported in part by the Marsha Rivkin foundation, the Ovarian Cancer Research Fund (OCRF), the Moffitt Lung Cancer SPORE Career Development Grant, the Bankhead-Coley, James & Esther King Biomedical Program, and NCI grant CA164147 to X.Z.

Footnotes

AUTHOR CONTRIBUTIONS

M.Z. and S.X. performed most of the experiments, each with equal contributions.

SUPPLEMENTAL INFORMATION

Supplemental Information includes one table, seven figures, and Supplemental Experimental Procedures and can be found with this article online at http://dx.doi.org/10.1016/j.molcel.2014.04.028.

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Supplementary Materials

Sup1
NIHMS629308-supplement-Sup1.pdf (992.4KB, pdf)
Sup2
NIHMS629308-supplement-Sup2.pdf (3.9MB, pdf)

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